Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Lamothe, G. T. Articles by Marugg, J. D. Search for Related Content PubMed PubMed Citation Articles by Lamothe, G. T. Articles by Marugg, J. D. Agricola Articles by Lamothe, G. T. Articles by Marugg, J. D.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, November 2003, p. 6541-6549, Vol. 69, No. 11
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.11.6541-6549.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Reverse Transcription-PCR Analysis of Bottled and Natural Mineral Waters for the Presence of Noroviruses

Gilbert Thierry Lamothe,^1* Thierry Putallaz,² Han Joosten,² and Joey D. Marugg²

Product Technology Centre, Nestlé Waters Management and Technology, F-88804 Vittel Cedex, France,¹ Nestlé Research Center, Vers-chez-les-Blanc, CH-1000 Lausanne 26, Switzerland²

Received 6 December 2002/ Accepted 17 August 2003

Top Abstract Introduction Materials and Methods Results Discussion References

 
A seminested reverse transcription-PCR method coupled to membrane filtration was optimized to investigate the presence of norovirus (NV) RNA sequences in bottled and natural mineral waters. The recovery of viral particles by filtration varied between 28 and 45%, while the limit of detection of the overall method ranged from 6 to 95 viral particles. The assay was broadly reactive, as shown by the successful detection of 27 different viral strains representing 12 common genotypes of NVs. A total of 718 bottled and natural mineral water samples were investigated, including 640 samples of finished, spring, and line products (mostly 1 to 1.5 liters), collected from 36 different water brands of various types and from diverse geographic origins over a 2-year period. In addition, 78 samples of larger volume (10 and 400 to 500 liters) and environmental swabs were investigated. From the 1,436 analyses that were performed for the detection of NVs belonging to genogroups I and II, 34 samples (2.44%) were presumptively positive by seminested RT-PCR. However, confirmation by DNA sequence analysis revealed that all presumptive positive results were either due to nonspecific amplification or to cross-contamination. In conclusion, these results do not provide any evidence for the presence of NV genome sequences in bottled waters.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Noroviruses (NVs [formerly known as Norwalk-like viruses]) play a major role in nonbacterial outbreaks of acute gastroenteritis (28) and are regarded as the main foodborne viruses. Most cases are caused by person-to-person spread and by consumption of shellfish or other contaminated food items. There have been relatively few outbreaks that have been attributed to contact with or drinking of contaminated water (11, 17, 19-22) despite the large degree of exposure of humans to water. Furthermore, in all of these cases, the origin of NVs or the route of contamination could clearly be identified. To the best of our knowledge, there is no documented epidemiological evidence that bottled water has ever served as a vehicle for spreading NVs. Nevertheless, it was recently reported that NV genome sequences could be detected in natural mineral waters sold in Switzerland and other countries (7, 8). Although it was clearly emphasized in those studies that detection of NV-specific sequences by itself does not imply that infectious virus particles are also present, the findings caused serious concern about the safety of bottled waters.

These results prompted us to screen for the presence of NV genomes in a number of brands of bottled water. For this purpose, we have used a previously described seminested reverse transcription (RT)-PCR procedure (15) that was also applied in the Swiss survey of bottled waters (7, 8). The different steps of the procedure were evaluated for their efficiency, and the sensitivity of the detection method was further optimized. Here we present results of a study performed on 718 samples, including finished, spring, and line products from 36 different brands of bottled and natural mineral waters. In addition, environmental factory samples (swabs) were investigated.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Virus strains and controls.
Clinical stool samples positive for NVs, provided by the RIVM (Bilthoven, The Netherlands), were used as reference material for NV detection and to assay the relative specificity of the procedure (Table 1). One positive and one negative control for each NV genogroup were included for each series of samples (i.e., four controls per series). For this purpose, 10 µl of stool samples of each genogroup (10^-3 to 10^-6 dilutions, depending on the viral titer of the different stools) was inoculated into 1 liter of sterile water. These spiked samples were treated in the same way as bottled water samples and used as controls for the whole procedure. Sterile water (1 liter) was used in all experiments as a negative control. In addition to these controls, a 10^-8 dilution of a synthetic RNA standard produced by in vitro transcription (iSt-ENV; Biosmart, Bern, Switzerland), which contains priming sites for the two genogroups of NVs (GI and GII), was used as an RT-PCR control, while nuclease-free water (Ambion) was used as negative control of RT-PCR.

View this table: [in this window] [in a new window]

TABLE 1. Specimens of clinical stool samples positive for NV used as positive controls in this study

Concentration of viral particles and RNA extraction.
Small- (1 to 1.5 liters, depending on the bottling size standard) and medium-size samples (10 liters) were vigorously shaken and filtered through a positively charged membrane (Zetapor filter membrane; CUNO, Inc., Saint-Herblain, France) according to a previously described procedure (15), with the following modifications: prefilters were omitted because of the low particle content of the bottled waters investigated, and virus elution was performed at 4°C for 3 h for optimal viral recovery. The eluate (4 ml) was further concentrated to 140 µl by ultrafiltration with Ultrafree Biomax 100 centrifugal units according to the recommendations of the manufacturer (Millipore, Le Mont sur Lausanne, Switzerland).

Large-scale samples were investigated by adaptation of an official procedure used for the concentration of enteroviruses from large water samples (3). In short, water from the production site upstream of the bottling line was filtered through a sterile glass wool column at a moderate flow rate (20 to 25 liters/h) for 20 h (corresponding to a final amount of 400 to 500 liters) to allow the viruses to adsorb onto the glass wool. After filtration, concentrated water samples were transported on ice to the laboratory for further processing. Viruses were desorbed from the glass wool by using three 100 ml samples of an elution buffer (50 mM glycin, 3% beef extract [pH 9.5]). The eluate was neutralized with 10 ml of 1 M HCl, and polyethylene glycol 6000 was added to a final concentration of 10%. The mixture was gently agitated for 1 h and incubated overnight at 4°C to precipitate viruses. The solution was centrifuged for 90 min at 10,000 x g at 4°C to collect the viruses, and the pellet was carefully resuspended in 8 ml of phosphate-buffered saline (PBS). Viral particles were further concentrated to 140 µl by ultrafiltration with Ultrafree Biomax 100 centrifugal units as described above.

RNA extraction and RT-PCR.
Viral RNA was extracted from retentates with the Qiaamp Viral RNA mini kit according to the manufacturer's instructions (Qiagen, Hilden, Germany).

The RT-PCR protocol was derived from a genogroup-specific assay previously described by Gilgen et al. (15). Prior to the RT the RNA was denatured by incubation at 70°C for 3 min directly followed by chilling on ice. The Sensiscript RT kit (Qiagen) was used to reverse transcribe 10 µl of viral RNA added to a master mix (10 µl) composed of 1x RT buffer, 400 µM each deoxynucleoside triphosphate (dNTP), 10 U of RNase inhibitor (RNasin; Promega, Madison, Wis.), 1 µM the appropriate oligonucleotide primer (Table 2), and 1 µl of Sensiscript reverse transcriptase and nuclease-free water (Ambion, Austin, Tex.). RT was achieved by incubation at 41°C for 1 h followed by 5 min at 95°C to inactivate the enzyme. The cDNA was subsequently amplified by seminested PCR, carried out in two steps (PCR1 and PCR2).

View this table: [in this window] [in a new window]

TABLE 2. Standard RT-PCR assay primers and conditions used in this study

PCR1 contained 20 µl of the cDNA mixed with 30 µl of a master mix containing 1x PCR buffer for Platinum Taq DNA polymerase (Invitrogen Life Technologies, Basel, Switzerland), 0.25 µM each primer (Table 2), MgCl₂ to the required final concentration (Table 2), nuclease-free water (Ambion), and 1 U of Platinum Taq DNA polymerase (Invitrogen Life Technologies). Thermal cycling was performed as follows: 2 min at 94°C for initial denaturation; 25 cycles of 20 s at 94°C, 45 s at 58°C, and 30 s at 68°C; and a final extension step of 3 min at 68°C.

PCR2 contained 1 µl of PCR1 mixed with 49 µl of a master mix containing 1x PCR buffer for Platinum Taq DNA polymerase (Invitrogen Life Technologies), 0.5 µM each primer (Table 2), MgCl₂ to the required final concentration (Table 2), nuclease-free water (Ambion), and 2 U of Platinum Taq DNA polymerase (Invitrogen Life Technologies). Thermal cycling was performed as follows: 2 min at 94°C for initial denaturation; 45 cycles of 20 s at 94°C, 45 s at 55°C, and 20 s at 68°C; and a final extension step of 3 min at 68°C.

Given that only 20% (10 µl out of 50) of the total RNA was tested for each sample in the RT-PCR system, it should be noted that results are representative of one-fifth of the volume of water sampled. This corresponds to 200 to 300 ml for small-scale samples, 2 liters for medium-scale samples, and 80 to 100 liters for large-scale samples.

Analysis of PCR products.
Twenty microliters of PCR2 was loaded on 3% low-melting-point agarose gels (1% SeaPlaque agarose and 2% NuSieve GTG agarose; BioWhittaker Molecular Applications, Verviers, Belgium). The gel was stained with ethidium bromide, and bands were visualized with a UV transilluminator. PCR fragments of the expected size for GI (241 bp) or GII (203 bp) were further analyzed by DNA sequencing.

PCR fragments were cloned prior to sequence analysis to ensure high-quality sequencing data. PCR products were first purified by using the Qiaquick PCR purification kit according to the manufacturer's instructions (Qiagen). Purified PCR fragments were ligated into the pCR-Blunt vector (Invitrogen Life Technologies) and transformed into chemically competent TOP10 Escherichia coli cells (Invitrogen Life Technologies) according to the recommendations of the manufacturer. Recombinant clones were screened by PCR with universal M13 (-40) forward and reverse primers. PCR products corresponding to positive clones were purified with the Qiaquick PCR purification kit (Qiagen). DNA sequencing of the two strands was either performed by Microsynth (Balgach, Switzerland) or done in house using the LiCor 4000 automatic sequencer (MWG-Biotech, Ebersberg, Germany). In the latter case, infrared dye (IRD41)-labeled primers were used in sequencing reactions, which were carried out with the Thermo Sequenase fluorescent-labeled primer cycle sequencing kit (Amersham Pharmacia Biotech, Amersham, United Kingdom). Sequences were assembled and aligned by using programs from the Lasergene package (DNAstar, Madison, Wis.). PCR sequences were compared to those in databases by using BLAST programs.

MPN-PCR analysis.
The viral titer for two strains of NVs (Valetta virus, GI; Hawaii virus, GII) present in stool samples was determined and expressed as PCR-amplifiable units (PCRU) per milliliter. Stools were thoroughly mixed with an equal volume of PBS and centrifuged at 5,000 x g for 10 min. The supernatant was collected, aliquoted, and used for RNA extraction. Ten-fold RNA dilutions were prepared and analyzed in triplicate by RT-PCR. The number of PCR positive samples seen on agarose gels for given dilution triplets were used to calculate statistical DNA template population estimates by direct most-probable-number PCR (MPN-PCR) according to Fode-Vaughan et al. (14). Calculations were made with Microsoft Excel spreadsheet and its Solver tool (Frontline Systems) as described previously (10). DNA population estimates derived from MPN-PCR values were converted to PCRU per milliliter) for each sample, taking into account all of the dilution steps along the procedure.

TaqMan-based quantitative PCR assays.
We developed two assays specific for a Valetta virus and a Hawaii virus commonly used as positive controls in our experiments (Table 1). The DNA sequences between primers SRI-1 and SRI-2 and between primers SRII-1 and SRII-2 for GI and GII, respectively, were determined for each viral strain and used as templates for the design of the required primers and probes. Primer Express software (version 1.0) and its guidelines were used for the primer probe design, together with guidelines from PE Applied Biosystems (Foster City, Calif.). Primers and probes (Table 3) were synthesized by Eurogentec (Herstal, Belgium). Amplification reaction mixtures (50 µl) contained 10 µl of cDNA prepared as described in the section on RNA extraction and RT-PCR, plus 1x TaqMan buffer, 0.2 µM virus-specific probe; and 0.3 µM each virus-specific forward and reverse primers. PCR samples and controls were prepared in triplicates. The reaction tubes were MicroAmp Optical tubes, together with MicroAmp optical caps. All consumables were supplied by PE Applied Biosystems. The amplification profile was as follows: 2 min at 50°C, followed by 10 min at 95°C and 45 cycles of 95°C for 15 s and 58°C for 1 min. Reactions were performed in the ABI Prism 7700 sequence detection system (PE Applied Biosystems) according to the instructions of the manufacturer. PCR products were detected directly by monitoring the increase in fluorescence from the dye-labeled virus-specific 5'-FAM, 3'-TAMRA TaqMan probe. Different amplifications were compared by their respective threshold cycles (C_T). The C_T values were plotted against log input and gave standard curves for quantitation of unknown samples and possibilities to estimate the amplification efficiency in the reaction. The limit of detection of the quantitative PCR assays was estimated by investigating 10-fold serial dilutions of known amounts of template DNA. The number of template molecules was determined by densitometry and standard protocols (32).

View this table: [in this window] [in a new window]

TABLE 3. Quantitative RT-PCR assay primers and conditions used in this study

Evaluation of the filtration procedure.
Filtration efficiency was determined by RNA quantitation of 10-fold serial dilutions (in the range of 0.2 x 10³ to 0.2 x 10⁷ PCRU) from a virus stock. For this, each viral dilution was either extracted directly or spiked into 1 liter of sterile water and filtered before extraction. The relative quantitation (filtered versus nonfiltered virus samples) allowed us to calculate the fraction of viruses recovered from the inoculum and to deduce the filtration efficiency. All assays were performed in triplicate for a virus of each genogroup (Valetta and Hawaii strains). Filtration and RNA extraction were performed as described for standard samples, and the quantitative PCR assays described above were applied to these samples after RT. The loss of viral RNA during the filtration step was calculated from the difference between C_T values of filtered samples and C_T values of nonfiltered ones (C_T). Consequently, the ratio of RNA concentration between two given C_T values is calculated as 2C_T. From this, the percent viral recovery from the filtration/elution step is deduced as 100/2C_T.

Experimental design and collection of water samples.
Water samples were obtained from 36 different brands of bottled waters. The waters were from diverse geographic origins (5 continents, 19 different countries) and included both carbonated and still waters with different features (i.e., aquifer type, mineral composition, and pH).

Finished products of bottled waters (brands E, S, X, and AF) were investigated for the presence of NV genomes on a weekly or biweekly basis in a long-term survey (up to a year). In addition, production samples including spring and line products taken at different stages of the bottling process were included in this survey (Table 4). Finished products of 32 other brands were investigated on a weekly basis (4 to 6 weeks in total) in a short-term survey (Table 4). All of the samples described above were investigated on a small-scale (1 to 1.5 liters) volume and were collected from various factories producing bottled water. Medium-scale (10 liters) water samples of finished products (brands E and X) were analyzed by the same filtration protocol (Table 4). Large-scale samples (400 to 500 liters) of finished products (brands X and AF) were collected and filtered through a glass wool column directly at the factory site, as described below.

View this table: [in this window] [in a new window]

TABLE 4. Samples investigated during the survey and presumptive results by RT-PCR

Environmental samples (swabs) were collected from eight different points representative of the bottling line in the factory producing brand S (Table 4). In short, borehole locations (S-7 to S-10), the storage tank inlet (S-11), the pump casing (S-12), and bottle cleaning and filling devices (S-13 and S-14) of this factory were monitored weekly over 2 months for the presence of NVs. Moistened cotton swabs were used to swab the 25- to 100-cm² sampled surfaces by a procedure similar to the one described by Taku et al. (35), who reported viral recovery in the range of 6 to 16%. Concentrated environmental samples were investigated by RT-PCR as for standard samples. All samples analyzed during this study were collected from December 1999 to December 2001.

Bacterial examinations.
All batches of water samples included in this work were investigated for the presence of several microorganisms in agreement with European regulation of natural mineral waters. In short, 250-ml water samples were investigated according to official validated procedures for the presence of coliforms (4), E. coli (4), intestinal enterococci (5), and Pseudomonas aeruginosa (2).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Performance assessment and optimization of the NV detection method.
Results in this study were generated with the protocol used by Beuret et al. (7, 8), which is based on a filtration step followed by a seminested RT-PCR procedure. In addition, the sensitivity and the relative specificity of the complete method with respect to NV sequence detection were evaluated and optimized in several ways.

The filtration efficiency was determined with 10-fold serial dilutions of a virus stock that were either directly quantified or spiked into sterile water and subjected to filtration. The filtration efficiency was assessed for the Valetta and Hawaii strains by means of a quantitative RT-PCR assay as described in Materials and Methods. The recovery was found independent of the sample volume in the range of 4 ml to 10 liters (data not shown). The results presented below correspond to 1-liter samples, which represent the majority of samples in this study. The loss of viral RNA during the filtration step was deduced from the difference between C_T values of virus dilutions obtained after filtration and C_T values of corresponding dilutions that were not subjected to filtration (C_T). The recovery after filtration and subsequent alkaline elution ranged from 28 to 35% (C_T = 1.82 to 1.53) for Valetta virus, depending on a low to high initial viral load, respectively (Fig. 1). Similarly, the recovery for Hawaii virus ranged from 30 to 45% (C_T = 1.76 to 1.16) (data not shown). In both cases, the viral recovery was higher for higher virus concentrations. The significant loss of viruses (55 to 72%) suggests that (i) filters do not retain all viral particles, (ii) viral elution from the filters is not complete, or (iii) degradation of viruses occurs due to the high pH of the elution buffer.

View larger version (13K): [in this window] [in a new window]

FIG. 1. 5'-nuclease PCR analysis of serial 10-fold dilutions of a Valetta virus variant strain inoculated into sterile, demineralized water. The amount of virus particles was analyzed by quantitative RT-PCR either after direct RNA extraction () or following the standard filtration/elution protocol (). CT values are plotted against the viral dilutions. The straight lines calculated by linear regression for direct RNA extraction (y = -3.61 x [virus] + 46.9) and for the standard filtration/elution protocol (y = -3.68 x [virus] + 49) show square regression coefficients of R2 = 0.999 (direct RNA extraction) and R2 = 0.996 (standard filtration/elution protocol). The linearity of the standard curves and the observation that the PCR operates with a constant efficiency confirm that the assay is well suited for quantitative measurements. Data points represent experiments performed in triplicate.

The RNA loss during the RNA extraction step was evaluated by the comparison of the quantitative RT-PCR signal of an RNA sample to the signal of the same sample extracted a second time. The quantitative RT-PCR signal intensities (C_Ts) were not significantly different (data not shown), and the loss during RNA extraction was therefore considered as negligible.

The sensitivity of our standard seminested RT-PCR procedure was compared to that of the quantitative RT-PCR assay in order to calculate the relative efficiencies of the two protocols. The comparison was made for the Valetta and Hawaii strains, and an MPN-PCR approach was used to evaluate the results in a statistical way (12). The viral titers expressed as PCRU per milliliter were estimated from the RNA population present in a sample by taking into account the respective dilution steps of each procedure. The values obtained from three independent sets of experiments for the seminested RT-PCR assay were very close to each other, indicating a good reproducibility for both genogroups (Table 5). The average titer obtained for the Valetta virus was 4.99 x 10⁹ PCRU/ml, as determined by seminested RT-PCR versus 1.11 x 10¹⁰ PCRU/ml by quantitative RT-PCR (Table 5), which corresponds to a 2.2-fold higher sensitivity for the specific quantitative assay. Results obtained for the Hawaii virus were 1.95 x 10⁷ PCRU/ml for the average titer (seminested RT-PCR) versus 1.11 x 10⁸ PCRU/ml by quantitative RT-PCR (Table 5), which corresponds to a 5.7-fold-higher sensitivity for the quantitative assay.

View this table: [in this window] [in a new window]

TABLE 5. Comparison of the efficiency of RT-PCR procedures for the determination of viral RNA titera

The overall limit of detection for the complete method was estimated for Valetta and Hawaii strains, taking into account (i) the relative efficiency of the seminested RT-PCR compared to the quantitative RT-PCR, (ii) the absolute limit of detection of the quantitative PCR assay, and (iii) the filtration efficiency for the respective virus strains. The absolute limit of detection of the quantitative PCR assay, determined by the use of a DNA standard of known concentration as described in the section on TaqMan-based quantitative PCR assays was found to be in the range of 1 to 5 template molecules for both assays (data not shown). Therefore, taking into account all of the parameters presented above (i, ii, and iii), the overall limit of detection for our procedure ranges from 6 to 39 particles for the Valetta strain and from 13 to 95 particles for the Hawaii strain.

The relative specificity of our RT-PCR procedure was tested on 27 different NV clinical stool samples chosen among strains most frequently identified in outbreaks of gastroenteritis in The Netherlands (36). These specimens were representative of both NV GI and GII and of 12 different genotypes (Table 1). All stools were found positive, which demonstrated that the assay was broadly reactive.

The presence of RT-PCR inhibitors was evaluated for all brands of bottled waters included in this survey. One liter of each water was analyzed by using the standard protocol. Another portion of 1 liter was spiked with 10 µl of a dilution of a stool sample positive for GI and analyzed. Finally, a small amount of an RNA standard (300 PCRU) was added to the RNA extracts of all of these samples and subsequently submitted to RT-PCR. All of the spiked water samples were found positive by RT-PCR. In contrast, all control water samples (unspiked) were found negative. Moreover, the RNA standard was successfully detected in all of these samples. These results strongly suggest that inhibitors did not play a role in this study.

Analysis of bottled water samples.
The long-term study (Table 4) aimed at evaluating any seasonal impact on the water quality and to monitor the water from the spring to the bottle. In addition, environmental samples were investigated to evaluate the possible presence of NV sequences inside factory S. Altogether, the presence of NV genome sequence was investigated in 509 samples from the four brands. The vast majority (93.91%) of the samples were found negative for the presence of NV genome sequences by seminested RT-PCR (Table 4). Thirty-one samples were found presumptive positive for GI (12), GII (18), or both (1), since bands of the expected size were visible on agarose gels. The cloning and DNA sequence analysis of amplicons revealed that 10 PCR fragments were not of viral origin, and therefore had to be considered as nonspecific amplifications (Table 6). The other 22 PCR fragments were identical to positive controls used in the experiments and for that reason classified as cross-contaminations (Table 6). Furthermore, the distribution of the latter strictly followed the pattern of the stools used as positive controls during the survey. Independently, all presumptive positive samples for GI or GII were reinvestigated by RT-PCR using the RNA leftover from the extraction procedure. Fragments classified as nonspecific could generally be reamplified (8 out of 10), while those classified as cross-contaminations could not.

View this table: [in this window] [in a new window]

TABLE 6. Sequence analysis of the presumptive positive PCR fragments

A second, short-term survey included a wide range of bottled waters with diverse characteristics and included a total of 179 finished product samples from 32 other brands. These waters were monitored over a short period of time (4 to 6 weeks on average) in order to search for the presence of NV genome sequences (Table 4). This worldwide survey was conducted to establish the influence of the water source. Again, in this survey, the majority of the samples (99.44%) were found negative for the presence of NV genomes. One sample was found presumptive positive for GI by seminested RT-PCR, but its DNA sequence analysis showed that this PCR fragment had been generated by nonspecific amplification (Table 6).

Analysis of larger volumes of water.
We have investigated larger volumes of water in order to find out whether NV genome sequences could be present at very low concentrations. For this purpose, large water samples of finished products (10 liters for brands E and X and 400 to 500 liters for brands X and AF) were analyzed (Table 4).

From the 28 samples analyzed on a 10-liter scale for brands E and X, 26 were found negative for the presence of NV genome. Two samples (7.14%) were found presumptive positive for GI by RT-PCR and were classified as cross-contamination, since their DNA sequence was identical to the positive control (Table 6). These two samples were reinvestigated by using the RNA left over from the extraction procedure, but the repetition experiment did not generate any PCR fragment. The two samples of large volume (400 liters, brand AF; 500 liters, brand X) analyzed were both found negative.

Bacterial examinations.
All batches of bottled water included in this work that were investigated for the presence of NVs were also investigated for the presence of coliforms, Escherichia coli, intestinal enterococci, and P. aeruginosa in 250-ml samples. These results comply with European regulations on natural mineral waters, since all were found negative.

Top Abstract Introduction Materials and Methods Results Discussion References

 
We have used an optimized seminested RT-PCR technique coupled to membrane filtration to investigate the alleged presence of NV genome sequences in natural mineral waters (7, 8). The RT-PCR assay is broadly reactive, as shown by its ability to detect 27 different NV strains representative of at least 12 different genotypes from both genogroups. A 28 to 45% recovery of virus particles was achieved from water by membrane filtration and an alkaline elution procedure with an estimated detection limit of 6 to 95 genome equivalents after RT-PCR. These values are estimations specific for the Valetta and Hawaii viruses and may vary for other NV strains, depending on the ability of each set of degenerated primers to anneal on a given viral sequence.

Using this combined assay, all 718 samples investigated representing 36 different brands and comprising 1,436 analyses for NV GI and GII were confirmed negative. Samples were investigated on a small scale (1 to 1.5 liters) and included finished, spring, and line products, as well as swabs. In addition, larger volumes of water were also investigated (10 and 400 to 500 liters).

Taking into account that RT-PCR inhibitors did not play a role in our study, these results indicate that NV genome sequences were not present in the investigated natural mineral and bottled waters. Overall, our findings are in sharp contrast to results of two separate surveys published recently by a Swiss laboratory in which 33% of the tested bottled waters had been found positive for the presence of NV genome sequences (7, 8). Although these studies reported on a limited number of samples (n = 153 [7] and n = 63 [8]) and different brands were investigated, there are no obvious technical reasons to explain the large discrepancy between the results of their investigations and those presented here. In fact, the methodologies are very similar, including in both cases an identical membrane filtration step coupled to an RT-PCR procedure (seminested) using the same NV-specific primer sets (16). However, a remarkable difference between the applied methodologies was the apparent use of sense primers in the RT step by the Swiss cantonal laboratory (7), which in fact targets DNA negative-strand templates rather than viral positive RNA strands.

Natural mineral waters originate from deep groundwater reservoirs that are well protected from fecal contamination, as shown by the absence of fecal indicator organisms in both the aquifers and finished products during routine microbiological quality controls (24). In particular, all batches of the brands of bottled water analyzed in this study were found free of coliforms, E. coli, intestinal enterococci, and P. aeruginosa by routine investigations based on validated standard procedures. In addition, our findings are strongly supported by the absence of any published epidemiological link between the consumption of bottled waters and the occurrence of gastroenteritis cases or outbreaks. According to the scientific literature, cases in which drinking water had been found positive for NVs mainly concerned surface waters that were contaminated by a sewage discharge event (11, 19, 21). Other cases involved insufficiently protected wells where the geological context explained the route of fecal contamination (22). Additionally, major defects in water supply systems or pumps have been reported (17). In all of these cases, coliforms were detected in the water, confirming the occurrence of fecal contamination (11, 17, 19, 21, 22). However, one cannot exclude the possibility that NVs could be found in water in the absence of fecal indicators.

The application of RT-PCR for the analysis of environmental samples has been very useful to associate NVs with outbreaks of nonbacterial gastroenteritis (13, 34). The use of RT-PCR for monitoring water quality with respect to NVs has been proposed by different investigators (18, 26, 34), but several limiting factors need to be considered. First of all, the detection of viral genomes in water by standard RT-PCR methods does not provide information about the levels or infectivity of the viruses in question, which impedes a meaningful risk evaluation in case positive results are obtained. Second, the high sensitivity of RT-PCR is likely to contribute to PCR artifacts (cross- and carryover contamination, nonspecific amplification). Several multicenter studies for virus detection reported false-positive levels of 2, 13, and 12.6%, respectively, for similar PCR or RT-PCR assays (25, 29, 31). In this study, false-positive results were obtained in 35 of 1,436 reactions (2.44%, of which 1.67% were cross-contaminations and 0.77% were nonspecific amplifications), which is quite low when compared with false-positive levels reported for these multicenter studies. Nonspecifically amplified DNA can easily be generated, because low annealing temperatures are used during PCR to tolerate some mismatches between primers and NV templates. This is considered essential for successful generic detection of such a genetically diverse group of viruses (38). The nonspecific amplifications observed in this study can also be explained by the presence of large amounts of bacterial RNA coming from the water concentrate or from the carrier RNA used in the RNA extraction kit. Because the occurrence of a few false-positive results may be regarded as almost inevitable in these extremely sensitive assays, it is paramount to perform additional DNA sequence analysis to determine the origin of amplicons (1, 6, 27, 30, 33, 39; this work). We observed that direct sequencing performed repeatedly on a single PCR fragment with degenerate primers led to numerous sequencing ambiguities and errors in the range of 4 to 8% over 200 bp (data not shown). It is therefore essential to clone each PCR fragment prior to sequence determination as well as to perform double-stranded sequencing to get reliable sequence information. Nested RT-PCR formats are considered to be less favorable for NV detection because of the higher risk of cross-contamination in diagnostic laboratories (38). One can expect the use of a single PCR step instead of a nested or seminested procedure to lower the risk of contamination in future studies. The last points important for the general acceptance of RT-PCR for routine monitoring of NVs are the validation and standardization of the methods currently in use in various laboratories to demonstrate the reliability, sensitivity, and ruggedness of the technique. These points have been shown to be very difficult to reach in several multicenter studies (25, 29, 31, 38). In conclusion, RT-PCR remains very laborious, expensive, and restricted to specialized laboratories. Perhaps, the best alternative for risk assessment of water sources with respect to NVs would be to use a sensitive cell culture system. Unfortunately, all attempts to develop such an assay have been unsuccessful for the last 30 years. Furthermore, the monitoring of bacteriophages has been considered, given their general resistance and persistence in the environment together with the possibility to cultivate them in a simple manner. However, their application as reliable indicators of fecal pollution remains to date a controversial issue (9, 23). Therefore, in view of the results presented above, together with the absence of epidemiological data linking the consumption of natural mineral water to NV outbreaks, we believe that NVs are not a real issue for bottled water safety.

 
We thank Marion Koopmans and Ana Maria de Roda Husman for the clinical stool samples of NVs. We thank Louis Schwartzbrod and Christophe Gantzer for their help and advice on the setting up of a glass wool column technique for the concentration of viruses from large water samples. We also thank Rolf Meyer for critical review of the manuscript.

 
* Corresponding author: Mailing address: Nestlé Waters M.T., Product Technology Centre, B.P. 101, F-88804 Vittel Cedex, France. Phone: 33/3 29 08 77 28. Fax: 33/3 29 08 71 32. E-mail: Gilbert.Lamothe{at}waters.nestle.com.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Ando, T., S. S. Monroe, J. R. Gentsch, Q. Jin, D. C. Lewis, and R. I. Glass. 1995. Detection and differentiation of antigenically distinct small round-structured viruses (Norwalk-like viruses) by reverse transcription-PCR and Southern hybridization. J. Clin. Microbiol. 33:64-71.[Abstract]
2
2. Anonymous. 1989. Testing water—bacteriological examinations of swimming pool water. AFNOR norm procedure NF T 90-421. Association Française de Normalisation, Saint-Denis La Plaine, France.
3
3. Anonymous. 1996. Testing water—detection of enterovirus-method by concentration on glass wool and detection by cell culture. AFNOR norm procedure XP T 90-451. Association Française de Normalisation, Saint-Denis La Plaine, France.
4
4. Anonymous. 2000. Water quality—detection and enumeration of Escherichia coli and coliform bacteria. Part 1. Membrane filtration method. ISO 9308-1. International Organization for Standardization, Geneva, Switzerland.
5
5. Anonymous. 2000. Water quality—detection and enumeration of intestinal enterococci. Part 2. Membrane filtration method. ISO 7899-2. International Organization for Standardization, Geneva, Switzerland.
6
6. Atmar, R. L., F. H. Neill, J. L. Romalde, F. Le Guyader, C. M. Woodley, T. G. Metcalf, and M. K. Estes. 1995. Detection of Norwalk virus and hepatitis A virus in shellfish tissues with the PCR. Appl. Environ. Microbiol. 61:3014-3018.[Abstract]
7
7. Beuret, C., D. Kohler, A. Baumgartner, and T. M. Lüthi. 2002. Norwalk-like virus sequences in mineral waters: one-year monitoring of three brands. Appl. Environ. Microbiol. 68:1925-1931.[Abstract/Free Full Text]
8
8. Beuret, C., D. Kohler, and T. Luthi. 2000. Norwalk-like virus sequences detected by reverse transcription-polymerase chain reaction in mineral waters imported into or bottled in Switzerland. J. Food Prot. 63:1576-1582.[Medline]
9
9. Bosch, A. 1998. Human enteric viruses in the water environment: a minireview. Int. Microbiol. 1:191-196.[Medline]
10
10. Briones, A. M., Jr., and W. Reichardt. 1999. Estimating microbial population counts by 'most probable number" using Microsoft Excel. J. Microbiol. Methods 35:157-161.[CrossRef][Medline]
11
11. Brown, C. M., J. W. Cann, G. Simons, R. L. Fankhauser, W. Thomas, U. D. Parashar, and M. J. Lewis. 2001. Outbreak of Norwalk virus in a Caribbean island resort: application of molecular diagnostics to ascertain the vehicle of infection. Epidemiol. Infect. 126:425-432.[CrossRef][Medline]
12
12. De Man, J. C. 1975. The probability of most probable numbers. Eur. J. Appl. Microbiol. 1:67-78.
13
13. Fankhauser, R. L., J. S. Noel, S. S. Monroe, T. Ando, and R. I. Glass. 1998. Molecular epidemiology of "Norwalk-like viruses" in outbreaks of gastroenteritis in the United States. J. Infect. Dis. 178:1571-1578.[CrossRef][Medline]
14
14. Fode-Vaughan, K. A., C. F. Wimpee, C. C. Remsen, and M. L. Collins. 2001. Detection of bacteria in environmental samples by direct PCR without DNA extraction. BioTechniques 31:598-602.[Medline]
15
15. Gilgen, M., D. Germann, J. Luthy, and P. Hubner. 1997. Three-step isolation method for sensitive detection of enterovirus, rotavirus, hepatitis A virus, and small round structured viruses in water samples. Int. J. Food Microbiol. 37:189-199.[CrossRef][Medline]
16
16. Hafliger, D., M. Gilgen, J. Luthy, and P. Hubner. 1997. Seminested RT-PCR systems for small round structured viruses and detection of enteric viruses in seafood. Int. J. Food Microbiol. 37:27-36.[CrossRef][Medline]
17
17. Hafliger, D., P. Hubner, and J. Luthy. 2000. Outbreak of viral gastroenteritis due to sewage-contaminated drinking water. Int. J. Food Microbiol. 54:123-126.[CrossRef][Medline]
18
18. Huang, P. W., D. Laborde, V. R. Land, D. O. Matson, A. W. Smith, and X. Jiang. 2000. Concentration and detection of caliciviruses in water samples by reverse transcription-PCR. Appl. Environ. Microbiol. 66:4383-4388.[Abstract/Free Full Text]
19
19. Kaplan, J. E., R. A. Goodman, L. B. Schonberger, E. C. Lippy, and G. W. Gary. 1982. Gastroenteritis due to Norwalk virus: an outbreak associated with a municipal water system. J. Infect. Dis. 146:190-197.[Medline]
20
20. Kappus, K. D., J. S. Marks, R. C. Holman, J. K. Bryant, C. Baker, G. W. Gary, and H. B. Greenberg. 1982. An outbreak of Norwalk gastroenteritis associated with swimming in a pool and secondary person-to-person transmission. Am. J. Epidemiol. 116:834-839.[Abstract/Free Full Text]
21
21. Kukkula, M., L. Maunula, E. Silvennoinen, and C. H. Von Bonsdorff. 1999. Outbreak of viral gastroenteritis due to drinking water contaminated by Norwalk-like viruses. J. Infect. Dis. 180:1771-1776.[CrossRef][Medline]
22
22. Lawson, H. W., M. M. Braun, R. I. Glass, S. E. Stine, S. S. Monroe, H. K. Atrash, L. E. Lee, and S. J. Englender. 1991. Waterborne outbreak of Norwalk virus gastroenteritis at a southwest US resort: role of geological formations in contamination of well water. Lancet 337:1200-1204.[CrossRef][Medline]
23
23. Leclerc, H., S. Edberg, V. Pierzo, and J. M. Delattre. 2000. Bacteriophages as indicators of enteric viruses and public health risk in groundwaters. J. Appl. Microbiol. 88:5-21.[CrossRef][Medline]
24
24. Leclerc, H., and A. Moreau. 2002. Microbiological safety of natural mineral water. FEMS Microbiol. Rev. 26:207-222.[CrossRef][Medline]
25
25. Lina, B., B. Pozzetto, L. Andreoletti, E. Beguier, T. Bourlet, E. Dussaix, L. Grangeot-Keros, B. Gratacap-Cavallier, C. Henquell, M. C. Legrand-Quillien, A. Novillo, P. Palmer, J. Petitjean, K. Sandres, P. Dubreuil, H. Fleury, F. Freymuth, I. Leparc-Goffart, D. Hober, J. Izopet, H. Kopecka, Y. Lazizi, H. Lafeuille, P. Lebon, A. Roseto, E. Marchadier, B. Masquelier, B. Picard, J. Puel, J. M. Seigneurin, P. Wattre, and M. Aymard. 1996. Multicenter evaluation of a commercially available PCR assay for diagnosing enterovirus infection in a panel of cerebrospinal fluid specimens. J. Clin. Microbiol. 34:3002-3006.[Abstract]
26
26. Lodder, W. J., J. Vinjé, R. van De Heide, A. M. de Roda Husman, E. J. T. M. Leenen, and M. P. G. Koopmans. 1999. Molecular detection of Norwalk-like caliciviruses in sewage. Appl. Environ. Microbiol. 65:5624-5627.[Abstract/Free Full Text]
27
27. McGoldrick, A., E. Bensaude, G. Ibata, G. Sharp, and D. J. Paton. 1999. Closed one-tube reverse transcription nested polymerase chain reaction for the detection of pestiviral RNA with fluorescent probes. J. Virol. Methods 79:85-95.[CrossRef][Medline]
28
28. Metcalf, T. G., J. L. Melnick, and M. K. Estes. 1995. Environmental virology: from detection of virus in sewage and water by isolation to identification by molecular biology—a trip of over 50 years. Annu. Rev. Microbiol. 49:461-487.[Medline]
29
29. Muir, P., A. Ras, P. E. Klapper, G. M. Cleator, K. Korn, C. Aepinus, A. Fomsgaard, P. Palmer, A. Samuelsson, A. Tenorio, B. Weissbrich, and A. M. van Loon. 1999. Multicenter quality assessment of PCR methods for detection of enteroviruses. J. Clin. Microbiol. 37:1409-1414.[Abstract/Free Full Text]
30
30. Pellett, P. E., T. J. Spira, O. Bagasra, C. Boshoff, L. Corey, L. de Lellis, M. L. Huang, J.-C. Lin, S. Matthews, P. Monini, P. Rimessi, C. Sosa, C. Wood, and J. A. Stewart. 1999. Multicenter comparison of PCR assays for detection of human herpesvirus 8 DNA in semen. J. Clin. Microbiol. 37:1298-1301.[Abstract/Free Full Text]
31
31. Quint, W. G. V., R. A. Heijtink, J. Schirm, W. H. Gerlich, and H. G. M. Niesters. 1995. Reliability of methods for hepatitis B virus DNA detection. J. Clin. Microbiol. 33:225-228.[Abstract]
32
32. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
33
33. Schwab, K. J., F. H. Neill, M. K. Estes, T. G. Metcalf, and R. L. Atmar. 1998. Distribution of Norwalk virus within shellfish following bioaccumulation and subsequent depuration by detection using RT-PCR. J. Food Prot. 61:1674-1680.[Medline]
34
34. Schwab, K. J., F. H. Neill, F. F. Le Guyader, M. K. Estes, and R. L. Atmar. 2001. Development of a reverse transcription-PCR-DNA enzyme immunoassay for detection of "Norwalk-like" viruses and hepatitis A virus in stool and shellfish. Appl. Environ. Microbiol. 67:742-749.[Abstract/Free Full Text]
35
35. Taku, A., B. R. Gulati, P. B. Allwood, K. Palazzi, C. W. Hedberg, and S. M. Goyal. 2002. Concentration and detection of caliciviruses from food contact surfaces. J. Food Prot. 65:999-1004.[Medline]
36
36. Vinje, J., J. Green, D. C. Lewis, C. I. Gallimore, D. W. Brown, and M. P. Koopmans. 2000. Genetic polymorphism across regions of the three open reading frames of "Norwalk-like viruses." Arch. Virol. 145:223-241.[CrossRef][Medline]
37
37. Vinjé, J., and M. P. G. Koopmans. 2000. Simultaneous detection and genotyping of "Norwalk-like viruses" by oligonucleotide array in a reverse line blot hybridization format. J. Clin. Microbiol. 38:2595-2601.[Abstract/Free Full Text]
38
38. Vinjé, J., H. Vennema, L. Maunula, C.-H. von Bonsdorff, M. Hoehne, E. Schreier, A. Richards, J. Green, D. Brown, S. S. Beard, S. S. Monroe, E. de Bruin, L. Svensson, and M. P. G. Koopmans. 2003. International collaborative study to compare reverse transcriptase PCR assays for detection and genotyping of noroviruses. J. Clin. Microbiol. 41:1423-1433.[Abstract/Free Full Text]
39
39. Whelen, A. C., T. A. Felmlee, J. M. Hunt, D. L. Williams, G. D. Roberts, L. Stockman, and D. H. Persing. 1995. Direct genotypic detection of Mycobacterium tuberculosis rifampin resistance in clinical specimens by using single-tube heminested PCR. J. Clin. Microbiol. 33:556-561.[Abstract]

---

Applied and Environmental Microbiology, November 2003, p. 6541-6549, Vol. 69, No. 11
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.11.6541-6549.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Poschetto, L. F., Ike, A., Papp, T., Mohn, U., Bohm, R., Marschang, R. E. (2007). Comparison of the Sensitivities of Noroviruses and Feline Calicivirus to Chemical Disinfection under Field-Like Conditions. Appl. Environ. Microbiol. 73: 5494-5500 [Abstract] [Full Text]  
• Butot, S., Putallaz, T., Croquet, C., Lamothe, G., Meyer, R., Joosten, H., Sanchez, G. (2007). Attachment of Enteric Viruses to Bottles. Appl. Environ. Microbiol. 73: 5104-5110 [Abstract] [Full Text]  
• Butot, S., Putallaz, T., Sanchez, G. (2007). Procedure for Rapid Concentration and Detection of Enteric Viruses from Berries and Vegetables. Appl. Environ. Microbiol. 73: 186-192 [Abstract] [Full Text]  
• Jothikumar, N., Lowther, J. A., Henshilwood, K., Lees, D. N., Hill, V. R., Vinje, J. (2005). Rapid and Sensitive Detection of Noroviruses by Using TaqMan-Based One-Step Reverse Transcription-PCR Assays and Application to Naturally Contaminated Shellfish Samples. Appl. Environ. Microbiol. 71: 1870-1875 [Abstract] [Full Text]  
• Sanchez, G., Joosten, H., Meyer, R., Beuret, C. (2005). Presence of Norovirus Sequences in Bottled Waters Is Questionable. Appl. Environ. Microbiol. 71: 2203-2205 [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Lamothe, G. T. Articles by Marugg, J. D. Search for Related Content PubMed PubMed Citation Articles by Lamothe, G. T. Articles by Marugg, J. D. Agricola Articles by Lamothe, G. T. Articles by Marugg, J. D.